Summary

Fe oxidation is one of Earth’s major biogeochemical processes, key to weathering, soil formation, water quality, and corrosion. However, our ability to track the contributions of Fe-oxidizing microbes is limited by our relatively incomplete knowledge of microbial Fe oxidation mechanisms, particularly in neutrophilic Fe-oxidizers. The genomes of many Fe-oxidizers encode homologs to an outer-membrane cytochrome (Cyc2) that has been shown to oxidize Fe in two acidophiles. Here, we demonstrate the Fe oxidase function of a heterologously expressed Cyc2 homolog derived from a neutrophilic Fe oxidizer. Phylogenetic analyses show that Cyc2 from neutrophiles cluster together, suggesting a common function. Sequence analysis and modeling reveal the entire Cyc2 family is defined by a unique structure, a fused cytochromeporin, consistent with Fe oxidation on the outer membrane, preventing internal Fe oxide encrustation. Metatranscriptomes from Fe-oxidizing environments show exceptionally high expression of cyc2, supporting its environmental role in Fe oxidation. Together, these results provide evidence that cyc2 encodes Fe oxidases in diverse Fe-oxidizers and therefore can be used to recognize microbial Fe oxidation. The presence of cyc2 in 897 genomes suggests that microbial Fe oxidation may be a widespread metabolism.

Introduction

Fe oxidation occurs in virtually all near-surface environments, producing highly reactive Fe oxyhydroxides that often control the fate of carbon, phosphorous, and other metals (Borch et al., 2010). It is commonly assumed that abiotic mechanisms are sufficient to account for Fe oxidation, particularly at near-neutral pH. However, Fe-oxidizing microbes are increasingly observed in a wide range of environments (Emerson et al., 2010; Kappler et al., 2015), leading us to ask to what extent microbes drive Fe oxidation. To address this, we need to confidently identify the Fe oxidase. But unlike other major microbial metabolisms, we have relatively incomplete knowledge of Fe oxidation pathways (Bird et al., 2011; Hedrich et al., 2011; Ilbert and Bonnefoy, 2013), and no candidates for a broadly distributed Fe oxidasehave emerged until now.

To prove this, we need functional information on Cyc2 from neutrophilic chemolithotrophic Fe-oxidizing bacteria (FeOB). Dark neutral pH environments are prevalent, and to date, these FeOB have been found in a wide variety of marine, terrestrial, and engineered environments, including aquifers, soils, sediments, hydrothermal vents, and water treatment systems (Kappler et al., 2015; Emerson and de Vet, 2015). In these environments, FeOB grow by coupling Fe oxidation to the reduction of O2 or nitrate, using this energy to fuel carbon fixation, thus serving as primary producers (Emerson et al., 2010). Known neutrophilic chemolithoautotrophic FeOB mostly fall within the marine Zetaproteobacteria (Mariprofundus spp., Ghiorsea spp.) and freshwater Betaproteobacteria (Gallionellales genera Gallionella, Sideroxydans, and Ferriphaselus) (Emerson et al., 2010; Kato et al., 2015; Mori et al., 2017). All sequenced genomes of Zetaproteobacteria and Gallionellales FeOB have cyc2 homologs, and these sequences are among the closest homologs to one another despite being from separate classes of Proteobacteria, forming a cluster separate from the acidophilic cyc2 (Kato et al., 2015; He et al., 2017). A second potential Fe oxidase gene, mtoA, was found in the Gallionellales Sideroxydans lithotrophicus ES-1 (Emerson et al., 2013), and functional and genetic information supports the role of MtoA and its homolog PioA in Fe oxidation (Jiao and Newman, 2007; Liu et al., 2012). However, few other FeOB genomes contain mtoA, suggesting that Cyc2 is potentially a more widespread Fe oxidase.

Thus, we set out to demonstrate the function of a Cyc2 from a neutrophilic Fe-oxidizer. We first analyzed the Cyc2 family phylogeny and then made structure-function predictions, which informed the design of the gene constructs that we expressed in E. coli. To support the Fe oxidase function, we performed whole cell Fe oxidation assays on Cyc2-expressing E. coli. To determine the environmental relevance, we analyzed cyc2 expression in a new marine Fe mat metatranscriptome and reanalyzed a published Fe-rich aquifer metatranscriptome (Jewell et al., 2016). Finally, we compare the genomic distribution and expression of cyc2 and mtoA, to better understand the relative significance of these two putative Fe oxidases.

Phylogeny of Cyc2

We started by producing a comprehensive phylogenetic tree of Cyc2 sequences acquired from databases (National Center for Biotechnology Information (NCBI) and Integrated Microbial Genomes (IMG)). To ensure that we were analyzing true homologs, we screened the sequences for appropriate length (352 to 587 aa, average/median 446 aa), the cytochrome c binding motif CXXCH, and beta barrel porin portion (see Cyc2 structure and conservation below). Our search yielded 897 unique near-full length sequences, which were reduced to 530 sequences when closely-related sequences were removed. The resulting Cyc2 fell into three distinct clusters, with sequences distributed amongst various bacterial taxa, largely Proteobacteria (Fig. 1, Supplemental File 1). Cyc2 homologs are present in all well-established neutrophilic microaerophilic chemolithotrophic FeOB (Table 1), many of which are obligate FeOB that lack other apparent Fe oxidase candidates. These microaerophilic FeOB Cyc2 sequences form a well-supported cluster (Cluster 1 in Fig. 1), with the marine Zetaproteobacteria within one subcluster, and the freshwater Gallionellales forming a separate subcluster that also includes the neutrophilic photoferrotrophic Chlorobi. This clustering suggests a common function for the Cyc2 in these neutrophilic FeOB.

Beyond neutrophilic FeOB, cyc2 is found in the genomes of various acidophilic FeOB, which suggests a common adaptation to Fe oxidation. There are homologs in various acidophilic FeOB genomes: Ferrovum spp., Thiomonas spp., and Burkholderiales GJ-E10, in addition to the functionally-verified Cyc2 from A. ferrooxidans and L. ferriphilum. Unlike the neutrophiles, the Cyc2 sequences from acidophiles do not form a single cluster, and instead are scattered across the Cyc2 tree. Notably, the two functionally verified Cyc2 from A. ferrooxidans and L. ferriphilum fall in different regions of the tree, Clusters 2 and 3 respectively. This presents the intriguing possibility that many or all of the Cyc2 homologs are Fe oxidases.

Cyc2 structure and conservation

To better understand the potential role of Cyc2 homologs and to prepare for functional studies, we performed sequence and structure predictions, focusing on Cyc2 from FeOB. Despite the great sequence diversity, the FeOB Cyc2 are all predicted to have a unique structure, a fused cytochrome-porin. All contain a signal sequence, a c-type cytochrome, and a porin (Supplemental Fig. 2). The signal sequence was predicted by SignalP, indicating that the protein is exported to the periplasm. The cytochrome portion is identifiable by a single CXXCH heme-binding motif, and is by far the most conserved part of the sequence (Fig. 2; Supplemental Fig. 2, 3). Conserved residues include a AXPXFAR[Q/K][T/Y] motif located 5 amino acids upstream of the CXXCH heme binding site (AXPXFARQT in Clusters 1 and 2 sequences; AXPXFARKY in Cluster 3). There is also a PXL motif 4 amino acids downstream of the CXXCH. This PXL motif can be found in many other cytochromes, such as the structurally characterized MtoD (Beckwith et al., 2015) and Cyc1 gene in Acidithiobacillus ferrooxidans (CYC41 in (Abergel et al., 2003)); the proline and lysine appear to help stabilize the heme (Abergel et al., 2003). In contrast, the AXPXFAR[Q/K][T/Y] motif is unique to Cyc2, and therefore could be used to distinguish Cyc2-like outer membrane cytochromes.

Alignment of cytochrome-containing section for Cyc2 from representative neutrophilic and acidophilic FeOB. Red=CXXCH heme-binding motif. Blue=other motifs discussed in the text. This section is preceded by a signal sequence, as detected by SignalP, and followed by a beta strand predicted by PSIPRED. See Supplemental Figure 2 for full Cyc2 alignment.

Two views of the Cyc2 porin model, generated using iTasser. The Cyc2 cytochrome (orange sphere=hypothesized location) is connected to the N terminal end of the porin (blue strand).

Cyc2 heterologous expression and functional assay

Cyc2 of neutrophiles are in a distinct cluster from functionally-characterized Cyc2 homologs, so its function still requires experimental evidence. Neutrophilic FeOB are challenging to grow in quantities sufficient for protein assays, so we took a heterologous expression approach. The cyc2 gene sequence from Mariprofundus ferrooxydans PV-1 (cyc2PV-1) was prepared for expression in E. coli by (1) codon optimization, (2) replacing the signal sequence with that of the E. coli outer membrane protein OmpA, and (3) adding a StrepII-tag at the C-terminus (Supplemental Fig. 6A). The resulting sequence was synthesized, cloned into pMAL-p4X, transformed into E. coli C43(DE3), and co-expressed with the pEC86 plasmid containing the ccm cytochrome c maturation genes under a constitutive promoter, to ensure proper cytochrome maturation under aerobic conditions (Arslan et al., 1998). The protein appeared to be somewhat toxic to E. coli, as the yield of Cyc2-expressing cells (OD600 = 1.1) was much lower than that of cells with empty vectors (2x-diluted OD600 = 1.9) even at low IPTG concentrations of 0.5 mM. Nevertheless, expression was successful, as shown by western blot using Strep-Tactin antibody against the protein N-terminal tag (Fig. 4A). This band runs close to the expected molecular weight of 43 kDa and contains heme, as established by heme-specific staining (Fig. 4B).

We tested the function by assaying Fe oxidation by Cyc2-expressing E. coli cells. Because of the low expression levels, we tested relatively dense cell suspensions (OD=2), washed and resuspended in fresh LB medium, buffered to pH 6 to help slow abiotic Fe oxidation. Fe(II) was added from an anoxic stock solution of FeCl2, to a concentration of 100 μM. The dense cell suspension appeared to stabilize Fe(II), as cells with an empty vector (i.e. plasmid without cyc2) showed considerably slower Fe oxidation relative to cell-free medium (Supplemental Fig 7A). Cyc2-expressing cells did indeed oxidize Fe(II): cells oxidized 41% of the Fe(II) within 2 minutes, and 73% within 10 min (Fig. 4C). On further addition of Fe(II) at 45 min, the Cyc2-expressing cells continued to oxidize Fe(II). In contrast, the empty vector control oxidized 14% of the Fe(II) in 10 min (Fig. 4C). Consistent results were obtained in triplicate experiments from cells taken from one specific expression stock (Fig. 4), as well as replicate assays using cells from different expressions (Supplemental Fig. 7B-D). Azide (3 mM) reduced Fe oxidation by 50%. Azide inhibits cytochromes by binding Fe in heme (Yoshikawa et al., 1998), suggesting that Cyc2 and/or the terminal cytochrome c oxidase were partially inhibited, though 3 mM azide may not have been sufficient to completely inhibit such a dense cell suspension. To confirm that Fe oxidation is due to the Cyc2 cytochrome, we expressed and assayed the porin portion of Cyc2PV-1 (Cyc2porin, 37 kDa as expected; Fig. 4A). These Cyc2porin-expressing cells oxidized Fe(II) very slowly (17% in 10 min; Fig. 4C), demonstrating that the cytochrome is required for Fe oxidation. Taken together, the data show that Cyc2-expressing E. coli accelerate Fe oxidation, so we conclude that Cyc2 confers the ability to oxidize Fe(II).

Expression of cyc2 in the environment

If Cyc2 is an Fe oxidase, we would expect high expression of cyc2 in Fe-oxidizing environments. To investigate this, we analyzed metatranscriptomes from two ecosystems dominated by Fe-oxidizing bacteria. We present a new dataset from the Loihi Seamount (Hawaii) hydrothermal vent Fe microbial mat and reanalyze an existing dataset from an Fe-rich alluvial aquifer in Rifle, Colorado (Jewell et al., 2016).

At the Loihi seamount, Fe-oxidizing microbial mats thrive where hydrothermal vents emit Fe(II)-rich fluids into oxygenated seawater (up to 700 μM Fe, <3 to 52 μM O2 in the mats (Glazer and Rouxel, 2009)). Here, Fe(II) is by far the most abundant electron donor, with relatively low or localized sulfide (Glazer and Rouxel, 2009). Using a remotely operated vehicle and syringe-based biomat sampler (Breier et al., 2012) containing RNA Later, we obtained a 17 mL sample of a surface mat for metagenomic and metatranscriptomic analyses. The microbial community was almost entirely Zetaproteobacteria (94.4% based on metagenome coverage), a class in which all cultured representatives are microaerophilic FeOB (n=15 (Emerson and Moyer, 2002; Emerson et al., 2007; McAllister et al., 2011; McBeth et al., 2011; Field et al., 2015; Makita et al., 2016; Mumford et al., 2016; Mori et al., 2017; Chiu et al., 2017; Laufer et al., 2017; Barco et al., 2017; Beam et al., in press)). One specific Zetaproteobacteria comprised 79.4% of the community. Overall, Zetaproteobacteria were also the most active, representing 90.5% of the mapped transcripts, with the dominant bin accounting for 78.9% of the transcripts. For this dominant Zetaproteobacteria bin, cyc2 is among the most highly expressed genes (91-99% percentile; Supplemental Fig. 8A). We looked for genes that indicated other possible respiratory metabolisms, but found none. This, combined with the high expression of cyc2, strongly suggests that cyc2 is a key gene in this marine Fe oxidation-based ecosystem.

In contrast to Zetaproteobacteria, freshwater Gallionellaceae genomes contain a wider range of electron transport genes: cyc2, mtoA, and in some cases the sulfur oxidation sox and reverse dsr genes. To see which were most highly expressed, we reanalyzed existing metatranscriptomes from an oxidation experiment at the well-studied Rifle aquifer. Situated next to the Colorado River, this aquifer was the subject of a long-term experimental study on uranium remediation by Fe- and S-reducing microbes (Williams et al., 2011). Acetate amendments resulted in a highly reduced zone with a large reservoir of Fe(II) minerals (Williams et al., 2009). Subsequently, Jewell et al. (2016) re-oxidized some of this Fe(II) by injecting nitrate-amended oxic groundwater, causing a bloom of Gallionellaceae. As the authors reported, cyc2 was among the most highly expressed genes, at 99.99-100th percentile in all three post-injection samples (Supplemental Fig. 8B). However, the cyc2 expression levels were not compared to mtoA, sox, and dsr genes, so their relative importance was unclear. Our re-analysis shows that the cyc2 gene was expressed at much higher levels than mtoA, sox genes, and dsr genes (Supplemental Table 2). In particular, cyc2 expression was approximately two orders of magnitude higher than mtoA (Table 2; Supplemental Fig. 9). This was true in individual bins when both were expressed, and also in total across all bins expressing these genes. Thus, while both putative Fe oxidases were expressed when Fe oxidation was stimulated, cyc2 was clearly preferentially expressed.

We can gain insight into the different niches of cyc2 and mtoA by examining the temporal expression patterns of different Gallionellaceae spp., (i.e. genomic bins; Supplemental Fig. 9). Four species/bins expressed both cyc2 and mtoA while >5 bins only expressed either cyc2 or mtoA. Overall, both putative Fe oxidase genes increased in expression level over time, but mtoA slightly peaked at the 3rd time point, when the aquifer was largely anoxic (as indicated by a low in terminal oxidase expression). The bin with the highest mtoA expression (22.6), showed peak mtoA expression during this anoxic period. This suggests that mtoA may be more useful under conditions of electron acceptor limitation.

The difference in cyc2 and mtoA expression at the Rifle site led us to ask whether cyc2 and mto/mtr homologs commonly co-occur in genomes (mtrABC encodes for an Fe reductase system homologous to the proposed MtoAB Fe oxidase system). We found that although cyc2 and mto genes co-occur in Gallionellaceae genomes, they are very rarely found in the same genome (Supplemental Fig. 10), suggesting that cyc2 and mto/mtr do indeed correspond to different niches.

Cyc2 homologs in other Fe-cycling and extracellular electron-transporting organisms

Of the 897 Cyc2 homologs analyzed (represented in Fig. 1, fully labeled tree in Supplemental File 1), very few are from genomes of well-established Fe-oxidizing taxa, in part because Fe oxidation is not typically tested in isolates. This brings up the question of whether or not all Cyc2 represent Fe oxidases. Although we do not know yet, we can gain insight from examining a few homologs in other organisms known to cycle Fe and/or engage in extracellular electron transport.

At least one of the organisms with cyc2, Dechloromonas aromatica RCB (Coates et al., 2001), is reported to be an anaerobic FeOB, coupling Fe oxidation with denitrification (Salinero et al., 2009). Nitrate-dependent Fe oxidation is controversial because these organisms often require organics, so it is not always clear if the Fe oxidation is enzymatic, or indirect via nitrite produced by heterotrophic denitrification (reviewed by Kappler et al. (2015)). We can now hypothesize that cyc2 encodes an Fe oxidase in D. aromatica, as well as some other denitrifiers not yet tested for Fe oxidation. If true, this would provide evidence in favor of enzymatic Fe oxidation in heterotrophs, and give a mechanism for studying this metabolism in isolates and the environment.

Another possibility is that Cyc2 functions more generally as a mechanism of extracellular electron transport (EET), i.e. to transfer electrons to and from a cell via the cell surface. This is not exclusive of Fe oxidation, as M. ferrooxydans PV-1 has been shown to oxidize a cathode (Summers et al., 2013) (though it is unknown whether Cyc2PV-1 is involved). Homologs of cyc2 are also present in the genomes of organisms known to conduct EET, but not proven to oxidize Fe. An example is the Gammaproteobacteria Tenderia electrophaga, which is the most active organism in a stable cathode-oxidizing consortia (Eddie et al., 2017). Curiously, T. electrophaga also has the conserved gene cassette with cycl and cytochrome c oxidases, found in FeOB (Supplemental Fig. 1), but this organism has not yet been isolated or shown to oxidize Fe. Homologs of cyc2 are also found in Geobacter spp., specifically G. bemidjiensis, G. uraniireducens, and Geobacter M18, M21, and Rf64, members of a clade that predominates in aquifers (Holmes et al., 2007; Merkley et al., 2015). These organisms reduce Fe(III), though it is thought that this happens via the Omc system. A study on the G. bemidjiensis proteome showed that the Cyc2 homolog Gbem_3353 was expressed at same levels during Fe(III) and fumarate reduction (Merkley et al., 2015), so function remained unclear, but could involve a more general EET role.

Conclusions

This study has provided multiple lines of evidence that Cyc2 is an Fe oxidase in diverse FeOB, including the first functional evidence that neutrophilic FeOB Cyc2 oxidizes Fe. Verifying the function of neutrophilic FeOB Cyc2 was important because this cluster of Cyc2 sequences is distinct and distant from the previously characterized Cyc2 and Cyt572 from acidophiles. It is still not clear if cyc2 is differentially expressed. If so, we would have a genetic marker of microbial Fe oxidation activity, which is otherwise difficult to distinguish from abiotic Fe oxidation at circumneutral pH. If not, the cyc2 gene is still a valuable way of recognizing the Fe oxidation potential in genomes and transcriptomes.

It is striking that there are so many Cyc2 homologs, including many from organisms not known to oxidize Fe. Phylogenetic and genomic analyses show that cyc2 has been horizontally transferred between known FeOB and with other organisms. The addition of Cyc2 alone conferred some Fe oxidation ability on E. coli, with Cyc2 likely interfacing with the existing electron transport system. This suggests that acquisition of cyc2 alone can allow an organism to oxidize Fe. If this is generally true, the abundance of Cyc2 homologs suggests that microbial Fe oxidation is more widespread than we currently recognize.

Experimental Procedures

Cyc2 phylogeny and amino acid identity calculations

Sequences with homology to Cyc2 were collected using blastp (Camacho et al., 2009) against the NCBI and IMG databases (maximum e-value 1*10-5). Query sequences were chosen to represent all major groupings of the Cyc2 tree, includingMariprofundus ferrooxydans PV-1, Gallionella capsiferriformans ES-2, Acidithiobacillus ferrooxidans ATCC 23270, Tenderia electrophaga, and Geobacter sp. FRC-32. To remove identical sequences, the resulting 2,413 sequences were clustered at 100% identity using CD-HIT, reducing the database to 977 cluster representatives (Li and Godzik, 2006). These sequences were imported into Geneious v.7.1.7, where they were aligned using MUSCLE (Edgar, 2004). The resulting alignment was used to filter out sequences of partial length, resulting in 897 sequences. Alignment columns with greater than 30% gaps were removed, and a maximum likelihood phylogenetic tree was built using RAxML (392 alignment columns, 100 bootstraps, CAT model of rate heterogeneity, JTT amino acid substitution model (Stamatakis, 2014)). From the resulting tree, sequences that were closely related and highly sampled (primarily from the Burkholderia and Xanthomonas genera) were removed, resulting in 530 full-length sequences that were re-aligned, and a final phylogenetic tree was built in RAxML (357 alignment columns, 300 bootstraps, CAT and JTT models). The resulting phylogenetic tree was colored and names customized using the Iroki program (Moore et al., 2017).

To calculate amino acid identities of the Cyc2 homologs, pairwise alignments between the 530 full-length Cyc2 sequences were constructed using Muscle (Edgar, 2004). Amino acid identity (AAI) values were calculated from these pairwise alignments based on the full-length sequence, as well as the cytochrome and porin domains separately. The cytochrome domain was defined as the conserved region starting 14 residues upstream from the heme binding site, and ending 21 residues downstream. The rest of the sequence downstream of this was defined as the β-barrel porin domain. The AAI values were imported into R and a histogram was plotted using the ggplot2 package (Wickham, 2009).

Structural modeling

Signal peptides were predicted using SignalP(Petersen et al., 2011) and secondary elements were predicted using PSIPRED (McGuffin et al., 2000). For identification of structural homologs to Cyc2, we uploaded sequences to the HHPRED tool, available as part of the Max Planck Institute Bioinformatics Toolkit (Söding et al., 2005). Information for the structural homologs are compiled in Supplementary Table 1. Structural modeling was carried out by HHpred/MODELLER (Söding et al., 2005; Webb and Sali, 2016), iTasser (Zhang, 2008), and Phyre (Kelley et al., 2015). The structural models from all three platforms were found to be in close agreement.

Cloning and heterologous expression of Cyc2

The sequence of cyc2 was optimized for expression in Escherichia coli, and the signal sequence replaced with the signal sequence of a native E. coli gene, ompA (sequences in Supplementalry Fig. 6). The gene was synthesized by Genscript (Piscataway, NJ, USA). The cyc2 gene was cloned into the EcoRI/ HindIII sites of the pMal-p4X plasmid (with the malE gene removed). This cyc2-containing plasmid was co-transformed into E. coli C43(DE3) with pEC86, a plasmid containing the cytochrome c maturation (ccm) genes under a constitutive promoter, to ensure heme insertion into Cyc2 (Arslan et al., 1998). We also co-transformed pEC86 and the pMalp4X plasmid without the cyc2 gene (and without malE) as an empty-vector control; cells with the empty vector control received the same treatment throughout heterologous expression and Fe-oxidation assays. For expression of cyc2, E. coli was grown aerobically at 37°C (with shaking at 200 RPM) in Lysogeny Broth (LB), buffered with 10 mM 2-(N-morpholino)ethanesulfonic acid (MES), pH 6; in addition, the medium also contained ampicillin (100 μg/mL) for propagation of the pMal-p4X plasmid, and chloramphenicol (30 μg/mL) for the pEC86 plasmid. After reaching mid-log phase (2.5 h), cultures were amended with 0.5 mM isopropyl β-D-1-thiogalactopyranoside (IPTG) for de-repression of the lac operon to induce cyc2 expression; induction proceeded for 20 hours at 18oC with shaking at ~200 RPM. Expression of the Cyc2porin was carried out in an identical manner, except for the absence of the pEC86 plasmid from cells expressing this construct, and therefore the absence of chloramphenicol from the media.

SDS-PAGE, western and heme staining

Cells were harvested and from cultures before IPTG induction and following 20 hours after induction. Biomass was adjusted to an optical density of 0.6 for the porin-only control and 1.2 for the empty-vector and Cyc2-expressing cells. Cells were lysed by re-suspension in 100 μL of 5X sodium dodecyl sulfate (SDS) running buffer (125 mM Tris, 1.25 M glycine, 0.5% SDS, pH 8.3) and passing the resuspension through a 27.5-gauge needle 10 times. Cells were then combined with gel loading buffer (50 mM Tris-HCl, 12.5 mM EDTA, 2% SDS, 10% glycerol, 0.02% bromophenol blue, pH 6.8), and centrifuged for 10 min at 15 000 x g. Fifteen μL of the sample was then loaded onto a 16% Tris-glycine SDS-PAGE gel, and ran at 100 V for 30 min, then 160 V for 40 min. The gel was then either Coomasie-stained (1 g Coomasie Brilliant Blue Stain in 10% acetic acid, 40% ethanol), or transferred to a PVDF membrane for heme stain and western blot. For heme and StrepII-tag detection, the SDS-PAGE gel was transferred to a PVDF membrane at 30 V for 16 h (4°C) in transfer buffer (25 mM Tris, 192 mM glycine, 20% methanol, pH 7.2). Heme peroxidase activity was assessed by washing the membrane with TBST buffer (20 mM Tris, 137 mM NaCl, 0.1% Tween-20, pH 7.6), and incubating for 30 minutes with Pierce ECL luminol and substrate (Carlson et al., 2013), before imaging on the Typhoon FLA 9500 (GE Healthcare Life Sciences). For StrepII-tag detection, the PVDF membrane was blocked for one hour with 5% bovine serum albumin (BSA) and 50 μg/mL avidin (from egg white) in TBST buffer, rinsed with TBST, and incubated for one hour with Precision Protein Streptactin-HRP conjugate (1:60,000 dilution, Biorad). The membrane was then washed with four 15-minute washes in TBST, then incubated with Pierce ECL luminol and substrate for 5 minutes before imaging on the Typhoon FLA 9500.

Fe oxidation assay

After induction of Cyc2 in E. coli, triplicate cultures were centrifuged at 3200 x g, and washed with sterile LB (supplemented with 10 mM MES, pH 6) before resuspension and incubation in 50 mL glass beakers in the same medium at an OD of 2 (final volume = 25 mL); these cell suspensions were stirred to ensure homogeneity and aeration. In the case of azide experiments, 75 μL of a fresh 1M sodium azide stock solution was added to 25 mL cell suspension (to a final concentration of 3 mM), and incubated for 5 min. In all experiments, FeCl2 was added from an anoxic, filter-sterilized 100 mM stock solution to a target concentration of 100 μM. Ferrozine measurements were taken over a period of one hour (ferrozine method adapted from (Stookey, 1970)). At each time point, a sample was taken from each of the triplicate cultures, and centrifuged at 15,000 x g for 20-30 s to remove cells. A 150 μL portion of the supernatant was then combined with 40 μL of 1.225 mM ferrozine, 50 μL of 6.87 M acetate buffer, and 10 μL of H2O inside 200 μL 96-well plates, and incubated for 15 min in the dark before absorption was measured at 562 nm using a plate reader (Perkin Elmer 1420 Multilabel Counter Victor3V). pH of the cell suspensions was monitored before and after each assay, and was found to be stable at pH 6 (within one-tenth of a pH unit).

Loihi Fe mat sampling and metatranscriptome analysis

Sampling

Sample J2-674-BM1-C123456 was collected from the Pohaku vents (Mkr 57) at Loihi Seamount, Hawaii using the Jason II remotely operated vehicle in 2013. Using the mat sampler designed by Breier et al. (2012), 60 mL of the top 1 cm of Fe mat was collected into each of six syringes, each pre-loaded with 50 mL of RNALater (Ambion, United States) for immediate DNA/RNA preservation. Upon recovery, samples were allowed to settle (~17 mL total mat material), overlying fluids were decanted, and samples were stored at -80°C until extraction.

Metagenome assembly and binning

Metagenomic reads were quality controlled (QC’ed) using trimmomatic and merged using FLASH (see pipeline: https://github.com/mooreryan/qc). QC’ed reads were then assembled using metaSPAdes (Nurk et al., 2017), with a k-mer sweep from 21 to 127. In total 82,765 contigs were produced, from 128 bp to 79,748 bp (average 1,203 bp). Contigs were then QC’ed so that only contigs with at least 1X coverage over 90% of their length were used. Contigs over 2,000 bp in length (10,352 total) were binned using Binsanity (Graham et al., 2017), CONCOCT (Alneberg et al., 2014), MaxBin (Wu et al., 2016), and MetaBAT (Kang et al., 2015). The best resulting bins were chosen using DAS Tool (Sieber et al., 2017). Bin completeness and redundancy were calculated using CheckM (Parks et al., 2015). The relative abundance of each bin was calculated using a length-normalized average of contig read coverage.

Manual bin curation

Loihi metagenome sample 674-BM1-C3 was dominated by a single Zetaproteobacteria OTU2 bin (S1_binsanity019), accounting for 79.4% of the total binned average read coverage (50.3% including unbinned contigs). This bin was estimated by CheckM to be 92.7% complete (3.03% redundancy, 0% strain heterogeneity), yet lacked any protein BLAST hits to cyc2. Because previous research has identified cyc2 within at least 9 genomic representatives from ZOTU2 (3 SAGs, 6 MAGs (Field et al., 2015; Fullerton et al., 2017)), we used two methods to find the cyc2 belonging to this bin: 1) subsampled reads for a simplified assembly and 2) used information from the assembly graph to locate cyc2 connected with this dominant bin. Quadruplicate, randomly-sampled read subsets at 10%, 2%, and 1% of the total number of quality-controlled reads were independently assembled and binned to simplify the assembly of the dominant ZOTU2 bin (starting at 1,352X coverage). Subsampling in this way allowed us to increase the quality of the dominant ZOTU2 bin, with a maximum completeness of 98.1% (2.27% redundancy, 0% strain heterogeneity), though unfortunately cyc2 was again not recovered from any of the binned contigs. However, the subsampled assembly was helpful in recovering cyc2 through use of the simplified assembly graph. Using blastp, 12 contigs were identified that had homology to Cyc2. All twelve of these contigs were found within one section of the assembly graph, connected through a minimal k-mer overlap of 21 bp to the assembly network containing the binned contigs of interest. No other bins were contained within this assembly graph network. Taking the section of the assembly graph containing these cyc2 contigs, overlap consensus assembly produced five unique contigs. Four of these contigs had sufficient length to confirm their clustering within the dominant ZOTU2 bin using VizBin (Laczny et al., 2015). These four contigs were combined with the dominant subsampled bin and used in further analysis.

Metatranscriptome recruitment and analysis

Metatranscriptome reads were recruited to metagenomic contigs using bowtie2 (Langmead and Salzberg, 2012). Expression estimates were calculated by dividing the total read count by the gene length and total sequencing effort (reads per thousand bp per million reads; RPKM). The relative abundance of the expression in each bin was calculated as the percent of total reads mapping to that bin, normalized to bin length. RPKM expression estimates from all expressed genes were then imported into R and plotted using the ggplot2 package (Wickham, 2009).

Nucleotide submission

Raw sequence data were submitted to the sequence read archive (SRA) at the National Center for Biotechnology Information (NCBI), with all appropriate metadata under project accession number PRJNA412510.

Reanalysis of Rifle aquifer metatranscriptome

To quantify cyc2 and other gene expression in an FeOB-dominated terrestrial ecosystem, we extracted information from the supplementary dataset of Jewell et al.(2016), a metatranscriptomic/metagenomic study of a nitrate/O2-amended ferruginous Rifle aquifer. This dataset consists of over 200,000 translated open reading frames (ORFs), grouped by bin and associated with gene expression data in units of RPKM (reads per thousand base pairs per million reads). From this dataset, amino acid sequences corresponding to ORFs in Gallionellaceae bins 22.1-22.9 were used as a protein sequence database, which we used for blastp to search for Cyc2, and Mto, Sox, Dsr protein sequences (Supplementary table 2).

RPKM expression values were then imported into R, log transformed, and plotted using the ggplot2 package.

cyc2 and mto/mtr gene distribution in genomes

To look for cyc2 and mtoAB/mtrABC gene distribution in genomes, we used blastp against the non-redundant NCBI database to search for homologs of these genes from a representative set of organisms: Sideroxydans lithotrophicus ES-1, Shewanella oneidensis MR-1, Rhodoferax ferrireducens T118, Magnetospirillum magneticum AMB-1, and Rhodopseudomonas palustris TIE-1. Blast results were analyzed using a custom Python script to identify which cyc2 and mto/mtr homologs co-occurred within the same genomes. Results were imported in R and plotted using the VennDiagram package.

Conserved gene cassette identification

We used a custom Python script to identify organisms that encode the conserved gene cassette described by Field et al. (2015). This cassette includes 2 subunits of the cbb3-type cytochrome c oxidase, cyc1, spermidine synthase, ferredoxin, and several other c-type cytochromes; we blasted these cassette genes against the non-redundant NCBI database (Release 84), and clustered close homologs according to gene ID, which allowed us to detect genomes that included genes of interest in close proximity, defined as within 20 genes of each other.

Acknowledgments

We thank Shawn Polson for assistance in bioinformatic analyses and the crews of R/V Thompson and ROV Jason for support during sampling at the Loihi Seamount. This research was funded by the National Science Foundation (EAR-1151682, OCE-1155290) and the Office of Naval Research (N00014-17-1-2640). The authors declare no conflict of interest.

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